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Forces and torques in the nucleus: chromatin under mechanical constraints1 Christophe Lavelle

Abstract: Genomic DNA in eukaryotic cells is organized in discrete chromosome territories, each consisting of a single huge hierarchically supercoiled nucleosomal fiber. Through dynamic changes in structure, resulting from chemical modifications and mechanical constraints imposed by numerous factors in vivo, chromatin plays a critical role in the regulation of DNA metabolism processes, including replication and transcription. Indeed, DNA-translocating enzymes, such as polymerases, produce physical constraints that chromatin has to overcome. Recent techniques, in particular single-molecule micromanipulation, have allowed precise quantization of forces and torques at work in the nucleus and have greatly improved our understanding of chromatin behavior under physiological mechanical constraints. These new biophysical approaches should enable us to build realistic mechanistic models and progressively specify the ad hoc and hazy ‘‘because of chromatin structure’’ argument often used to interpret experimental studies of biological function in the context of chromatin. Key words: DNA, nucleosome, chromatin, mechanical constraints, supercoiling, transcription. Re´sume´ : Dans les cellules eucaryotes, l’ADN ge´nomique est organise´ en territoires chromosomiques distincts, chacun e´tant compose´ d’une longue chaıˆne de nucle´osomes surenroule´e hie´rarchiquement. Par sa capacite´ a` changer dynamiquement de structure sous l’influence des modifications chimiques ou des contraintes me´caniques impose´es par de nombreux facteurs in vivo, la chromatine joue un roˆle crucial dans la re´gulation du me´tabolisme de l’ADN, notamment lors de la re´plication et de la transcription. En effet, des enzymes comme les polyme´rases produisent, en avanc¸ant le long de l’ADN, des contraintes physiques auxquelles la chromatine doit faire face. Des techniques re´centes, en particulier reposant sur la micromanipulation de mole´cules uniques, ont permis de mesurer pre´cise´ment les forces et couples en jeu dans le noyau et ont largement ame´liore´ notre connaissance du comportement de la chromatine soumise a` ces contraintes me´caniques physiologiques. Ces nouvelles approches biophysiques devraient nous permettre de construire des mode`les me´canistiques re´alistes et ainsi de pre´ciser l’argument ad hoc et flou, « a` cause de la structure de la chromatine », souvent invoque´ pour interpre´ter les e´tudes expe´rimentales de fonctions biologiques dans un contexte chromatinien. Mots-cle´s : ADN, nucle´osome, chromatine, contraintes me´caniques, surenroulement, transcription.

Introduction Chromatin is constantly changing its structure to dynamically and functionally accommodate DNA transcription, replication, recombination, and repair. Many reviews on chromatin structure and dynamics focus on the role played by the enzymes responsible for the chemical posttranslational modifications (PTMs) of histones and nucleosome remodeling. The aim of this paper is to add a physical dimension to the description of chromatin to provide a more complete framework in which the control of genetic expression and other DNA metabolism processes can be interpreted. Because thermodynamic and electrostatic aspects

have recently been extensively reviewed elsewhere (Korolev et al. 2007), we will focus on mechanical issues related to DNA transactions. Indeed, DNA-translocating enzymes, such as polymerases, produce physical constraints: as these molecular motors push, pull, and twist DNA, transient forces and torques develop within chromatin (Cozzarelli et al. 2006). The recent development of experimental tools allowing the precise application and measurement of pico-Newton (pN) forces, acting on single macromolecules, has opened new perspectives in biology (Bustamante et al. 2000; Clausen-Schaumann et al. 2000). This has been particularly striking in the chromatin field, where investigation of individual chromatin fibers has en-

Received 15 May 2008. Revision received 31 July 2008. Accepted 1 August 2008. Published on the NRC Research Press Web site at bcb.nrc.ca on 12 February 2009. C. Lavelle. Institut Curie, Centre de Recherche, Paris F-75248, France (e-mail: [email protected]); CNRS, UMR 168, Paris F-75248, France. 1This

paper is one of a selection of papers published in this Special Issue, entitled 29th Annual International Asilomar Chromatin and Chromosomes Conference, and has undergone the Journal’s usual peer review process.

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doi:10.1139/O08-123

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Since naked DNA almost never exists in eukaryotic nuclei, chromatin processing necessarily precedes DNA processing. It is now clear that chromatin, more than a natural barrier to DNA accessibility, has a critical regulatory role in DNA metabolism, effected through functional dynamic processes that have yet to be fully characterized. The socalled 30 nm fiber is the most common chromatin superstructure described and, therefore, appears to be a convenient organization level at which to study regulation processes (Lesne and Victor 2006). However, the precise structure, and even the existence, of such a discretely defined folding stage is still debated (van Holde and Zlatanova 2007). Obviously, chromatin is a continuous and polymorphic structure with successive and transient compaction states. While the existence of specific interchangeable distributions of chromatin structure along the genome (e.g., during differentiation) is usually referred to as chromatin polymorphism or plasticity, transient perturbations occurring at a shorter time-scale (e.g., during transcriptional activation) involve chromatin dynamics and elasticity (Ben-Haim et al. 2001). Chromatin resilience and flexibility describe the smooth response of chromatin to imposed mechanical and topological constraints (Bancaud et al. 2006). Now, knowing that chromatin is a plastic polymorphic dynamic elastic resilient flexible nucleoprotein complex, how well can we quantitatively characterize its physical properties? Obviously, recent micromanipulation tools are making this possible.

proaches, complemented by an array of high-resolution imaging techniques, including (cryo-)electron and atomic force microscopy. In typical experiments, one investigates DNA or nucleosomal templates one at a time by exerting a force and measuring the subsequent deformation of the substrate. In another type of experiment, DNA is used as a mechanosensor to measure protein interactions and enzymatic actions occurring on it. Hence, both the physiological substrate (DNA or chromatin fiber) by itself and its interaction with various proteins can be addressed. Single chromatin fiber assembly (Leuba et al. 2003) and response to tension (Bennink et al. 2001a; Brower-Toland et al. 2002; Cui and Bustamante 2000; Pope et al. 2005) or torsion (Bancaud et al. 2006, 2007) have revealed the existence of an internucleosomal attraction that maintains higher-order chromatin structure (Cui and Bustamante 2000), the multistep unwrapping of nucleosomal DNA (Brower-Toland et al. 2002; Pope et al. 2005), and the capability of chromatin to accommodate large amounts of torsional stress (Bancaud et al. 2006, 2007). Together, these approaches enhanced our understanding of chromatin dynamics and its response to various mechanical constraints. More particularly, 2 recent studies by Bancaud and coworkers (2006, 2007) have addressed, for the first time, the torsional response of single nucleosome arrays (Fig. 1c), revealing 2 surprising results. First, chromatin fibers show higher torsional resilience than naked DNA (Bancaud et al. 2006). This behavior has been explained by a dynamic equilibrium of 3 conformations of the nucleosome, corresponding to different crossing geometries adopted by DNA as it enters and exits the nucleosome, based on previous results from minicircle studies by Prunell and coworkers (De Lucia et al. 1999; Sivolob et al. 2003; Sivolob and Prunell 2004). Second, these chromatin fibers, after extensive positive supercoiling, display no nucleosome loss, but rather a hysteretic behavior in their mechanical response to torsion (Bancaud et al. 2007). This hysteresis was interpreted as a consequence of the trapping of positive turns in individual nucleosomes, through their transition to an altered form, called reversome (for reverse nucleosome), which can be related to the previously documented chiral transition of the tetrasome (Hamiche et al. 1996a). These results will be further discussed later.

Chromatin micromanipulation Biologically relevant forces vary from pN (thermal fluctuation, entropic forces), to tens of pN (produced by some powerful molecular motors), to hundreds of pN (noncovalent interactions, such as van der Waals, hydrogen, and ionic bonds), to thousands of pN (covalent bonds). Various tools have been developed that can exert and (or) measure such forces at the single-molecule level, mainly mechanical force transducers (atomic force microscope cantilevers, microneedles, optical fibers) and external field manipulators (photonic in optical tweezers, magnetic in magnetic tweezers, hydrodynamic in flow field systems) (Bustamante et al. 2003, 2000; Zlatanova and Leuba 2003) (Fig. 1). The study of bending, twisting, melting, and opening of DNA and chromatin have greatly benefited from these new ap-

DNA and chromatin mechanical parameters As mentioned before, chromatin fiber may conveniently be seen as a long flexible polymer hierarchically folded in the nuclear space. To better understand how this folding is accomplished, and more generally how chromatin fibers may respond to mechanical constraints, several elastic parameters have to be determined, including the bending rigidity, torsional rigidity, and stretching modulus. Micromanipulation approaches, coupled with modeling, have been crucial in estimating these parameters, both for DNA and for chromatin. However, one should keep in mind that, aside from divergences arising from experimental and data analysis differences, sequence effects (Lankas et al. 2000) and geometrical variability (Ben-Haim et al. 2001; Wedemann and Langowski 2002) can greatly influence, respectively,

abled remarkable progress in the understanding of this ubiquitous macromolecular nucleoprotein complex (Marko and Poirier 2003; Zlatanova and Leuba 2003). Although such biophysical approaches have also greatly improved our understanding of mitotic chromosome structure (Marko 2008), we will only consider dynamic processes occurring during interphase, with a particular emphasis on transcription. Both quantitative mechanical values (elastic parameters, forces and torques, structural transition energies) and qualitative issues (chromatin conformation, transcription dynamics) will be discussed. We hope this physical description will help in the understanding of the influence of mechanical constraints in the regulation of DNA metabolism in the nucleus.

Chromatin nanomechanics

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Fig. 1. Forces and torques assessment in vitro: single-molecule experiments. Various methods have been developed to manipulate single nucleosomal arrays. Depending on the set up, these experiments enable the study of nucleosome assembly and disassembly under constraints, or the application of force (tension) and (or) torque (torsion) to chromatin fibers to measure their mechanical response. These techniques use optical tweezers (a, b, e), magnetic tweezers (c, d), or flow (d, e, f) to apply constraints on one end of a chromatin fiber attached at the other end to the surface of a cover slip (a, c, d, e) or to the extremity of a micropipette (e, b). Chromatin can thus be pulled (a, b, c, d, e, f) or pulled and rotated (c). These various set-ups were used in the following studies: (a) Brower-Toland et al. (2005, 2002); (b) Bennink et al. (2001a), Cui and Bustamante (2000), Gemmen et al. (2005), Mihardja et al. (2006); (c) Bancaud et al. (2006, 2007); (d) Leuba et al. (2003); (e) Bennink et al. (2001b), Pope et al. (2005); and (f) Ladoux et al. (2000).

DNA and chromatin elastic parameters. Hence, the given values are mostly to be taken as averages (Tables 1 and 2). The persistence length, A, a measure of bending flexibility, has been determined for DNA in various experiments, leading to a consensus value of ~50 nm (150 bp) at physiological ionic strength (see, for instance, Hagerman 1988; Lu et al. 2001; and references therein). For chromatin, values are more divergent, from ~30–50 nm estimated from singlemolecule stretching (Bancaud et al. 2006; Bennink et al. 2001a; Cui and Bustamante 2000), recombination frequencies (Ringrose et al. 1999), and crosslinking probabilities (Dekker et al. 2002), to more than 200 nm estimated from recent in situ hybridization experiments (Bystricky et al. 2004). Remarkably, such different values are consistent with theoretical models, and could be explained by the polymorphic connection geometry between nucleosomes in the fiber (Ben-Haim et al. 2001; Wedemann and Langowski 2002). The torsional persistence length, C, a measure of twisting flexibility, has been estimated for naked DNA by various

bulk experiments, giving a value ~70 nm (200 bp) (Fujimoto and Schurr 1990; Shore and Baldwin 1983), consistent with single-molecule data (Strick et al. 1996) analyzed through an elastic model (Neukirch 2004), although higher values have also been reported (Byrant et al. 2003). The torsional persistence length of chromatin fiber, measured for the first time recently, has a surprising low value of 5 nm (Bancaud et al. 2006), which is almost 20 times lower than that of naked DNA, and is also lower than the value predicted by analytical models of chromatin fibers (Ben-Haim et al. 2001). As discussed earlier (see Chromatin and micromanipulation), however, this result is easily explained by conformational nucleosome dynamics, and is fully consistent with new chromatin models, including this nucleosomal property (Bancaud et al. 2006). The stretching modulus, s, a measure of stretching elasticity (i.e., the propensity of a polymer to extend under a given tension), is about 1100 pN for naked DNA, as estimated from single-molecule experiments (Cluzel et al. 1996; Smith et al. 1996; Wang et al. 1997). For chromatin fiber, a much Published by NRC Research Press

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Biochem. Cell Biol. Vol. 87, 2009 Table 1. Approximate elastic parameters. Structure DNA Chromatin fiber

Bending persistence length (nm) 50 30–200

Torsional persistence length (nm) 70–100 5

Stretching modulus (pN) 1100 5–150

Note: See the text for references.

Table 2. Approximate energy, force, and torque parameters. Phenomenom

Energy cost, force and torque

Energies Nucleosome conformational transition (shift between +/– linker DNAs crossing) Tetrasome chiral transition Nucleosome chiral transition (nucleosome–reversome transition) Nucleosome dissociation

2–4 kT 2 kT 6–8 kT (but 25 kT energy barrier) 20–40 kT

Force (pN) and torque (pN
nm) RNA polymerase (Escherichia coli) DNA polymerase (phage T7) DNA melting Chromatin fiber unfolding (compact 30 nm fiber to beads-on-a-string) Nucleosome disruption Chromatin response to torsion DNA response to torsion

25 pN/>5 pN
nm 40 pN 8 pN
nm <5 pN 20 pN/36 pN
nm (probably much less in vivo) <3 pN
nm <6 pN
nm

Note: See the text for references.

lower value of ~5–8 pN has been derived from the same kind of experiments (Bancaud et al. 2006; Cui and Bustamante 2000), which is close to the predicted value (BenHaim et al. 2001), and probably reflects the weak internucleosomal interactions disrupted during the mechanical extension of the fiber at low ionic strength. Confirming this view, the stretching modulus is indeed greatly enhanced in high salt conditions (Bennink et al. 2001a; Katritch et al. 2000). Two main features emerge from these studies. First, torsional and bending deformations propagate over about the same distance in DNA, whereas in chromatin, torsional flexibility may be as much as 40 times higher than bending flexibility. Second, the stretching modulus is much greater (up to 200 times) for DNA than for chromatin; hence, tension applied by a molecular motor on the DNA backbone will cause chromatin distortion well before DNA distortion.

ming from the existence of histone variants (Kamakaka and Biggins 2005), histone PTMs (Peterson and Laniel 2004), and sequence-dependent properties of the wrapped DNA (Sivolob et al. 2003). Furthermore, nucleosomes are in dynamic equilibrium within the chromatin fiber; they can partially unwrap or shift from an open state to a closed one, either positive or negative, depending on the orientation of exiting DNA strands (De Lucia et al. 1999). Hence, chromatin is not only a reserve of constrained negative supercoiling, with a mean of –1 supercoils stored per nucleosome (Prunell 1998), it may also act as a dynamic buffer, able to transiently absorb negative or positive torsional stress due, in particular, to the inherent conformational flexibility of nucleosomes (Bancaud et al. 2006; Sivolob et al. 2003). Part of the torsional stress could, alternatively, be dispersed as torsion in linker DNA. The question that remains is what balances these 2 possibilities in vivo.

Chromatin physical tuning

Tuning by nucleosome structural changes It has been suggested that positive torsional stress may trigger the conversion of nucleosomes into hexasomes or tetrasomes (nucleosomes lacking 1 or 2 H2A/H2B dimers, respectively), lexosomes (distorted nucleosomes), or reversomes (chirally reversed nucleosomes, discussed in Chromatin micromanipulation). These different conformational transitions have some energy cost (Sivolob et al. in press), and should therefore be favored by mechanical constraints. Roughly, it takes 2–4 kT (depending on the sequence of the wrapped DNA) to shift entry and exit nucleosomal DNA from a negative to a positive crossing. This value is roughly divided by 2 when linker histone is added, which brings the 2 DNA duplexes into parallel and facilitates their fluctuation (Sivolob and Prunell 2003). As for chiral transitions, it takes 2 kT for the tetrasome (left–right transition) and 6 kT for the nucleosome (nucleosome–reversome transition), although, in

If DNA can be considered a molecular spring with elastic constants given by its sequence, chromatin fiber can be regarded as a supramolecular metaspring, with elastic constants influenced not only by the DNA sequence wrapped in this structure, but also — as stated earlier (see DNA and chromatin mechanical parameters) — by the geometrical parameters of the fiber (individual nucleosome conformation, spacing between adjacent nucleosomes), the histone composition (histone PTM, histone variants), and the presence of linker histones and nonhistone proteins participating in chromatin fiber folding. All of these parameters may vary locally and dynamically, providing chromatin with a tunable elasticity critical for its function (Ben-Haim et al. 2001). Far from being a repetitive and static chromatin entity, each nucleosome has its own structural characteristics, stem-

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this last case, a rather high 25 kT initial barrier has to be reached (Bancaud et al. 2007). It is therefore likely that noncanonical nucleosomal structures will appear upon mechanical stress and participate in transcription regulation (Lavelle and Prunell 2007). Meanwhile, complete dissociation of the nucleosome requires ~20 kT, as measured from dilution methods on bulk chromatin (Ausio et al. 1984; Cotton and Hamkalo 1981), a value that has to be increased beyond 25 kT on some strong positioning sequences (Gottesfeld and Luger 2001, but see Thastrom et al. 2004 for a critical discussion on these results). Since nucleosomes do not represent a barrier to transcription elongation (Lavelle 2007), the question arises as to which mechanisms enable their destabilization in vivo. This point will be discussed in Transcription. Tuning by histone PTMs Chromatin carries numerous histone modifications, some of which may be inherited (making them truly epigenetics). Eight different types have been characterized to date, and their occurrence and function in various biological processes are regularly reviewed (see, for instance, Berger 2007; Kouzarides 2007). Two (nonexclusive) mechanisms have been proposed for the function of these modifications: first, a structural hypothesis, which supposes that chromatin packing may be altered directly (by a change in electrostatic charge, affecting the contact between adjacent nucleosomes or the interaction of histones with DNA), thus controlling the recruitment of DNA-binding proteins, such as transcription factors or repair enzymes; and second, a signaling hypothesis, which supposes that attached chemical moieties may alter the nucleosome surface to promote the association of chromatin-binding proteins. Strikingly, a single histone modification can modulate both higher-order chromatin structure and functional interactions with regulatory proteins (Shogren-Knaak et al. 2006; Shogren-Knaak and Peterson 2006), stressing the difficulty in deciphering the so-called histone code. Of all the known histone modifications, acetylation appears to be the most consensual one, favoring transcription (Allfrey et al. 1964; Hebbes et al. 1988; for reviews, see Turner 1991; Workman and Kingston 1998). Histone acetylation seems to influence transcriptional initiation, but not elongation (Roberge et al. 1991). This view is supported by studies showing that this modification is sometimes observed only on the flanking regions of genes (Tazi and Bird 1990; Turner et al. 1990). In the signaling scenario, acetylation promotes direct stimulation by changing the interaction of the histone tails with components of the transcriptional machinery (Pineiro et al. 1991). In the more favored structural scenario, acetylation enhances chromatin accessibility (Gorisch et al. 2005). The opening of chromatin upon histone acetylation could be due to a simple electrostatic effect, in which charge screening (acetylation neutralizes the positive charge of N-terminal lysine residues) induces nucleosome opening (De Lucia et al. 1999; Toth et al. 2006) and chromatin unfolding (Ausio 1992). Another suggestion is that histone acetylation potentially inhibits nucleosome interactions or alters the capacity of the H1 histone to form compact higher-order chromatin structures, such that active and (or) competent gene chromatin is maintained in a less

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folded state than the bulk of chromatin (Allan et al. 1982; Ridsdale et al. 1990). Tuning by histone variants Numerous histone nonallelic isoforms exhibit primary sequence differences in strategic regions of the histone fold (Bernstein and Hake 2006; Pusarla and Bhargava 2005). By providing nucleosomes with new structural properties, these histone variants participate in chromatin structural and functional polymorphisms. Indeed, several effects have been characterized, such as the partial unwrapping at the edges of H2A.Bbd nucleosomes (Bao et al. 2004; Doyen et al. 2006), the facilitated disassembly of CENP-A nucleosomes (Conde e Silva et al. 2007), the stabilization of H2A.Z nucleosomes (Park et al. 2004), and the destabilization of H3.3 nucleosomes (Jin and Felsenfeld 2007), which is enhanced by the simultaneous replacement of H2A with H2A.Z (Jin and Felsenfeld 2007). Furthermore, nucleosome assembly enzymes, known as histone chaperones, participate in both the assembly and disassembly of nucleosomes, enhancing chromatin plasticity and dynamics. These chaperones not only facilitate the deposition of canonical histones behind the replication fork in S phase, they also mediate the incorporation of histone variants in a replication-independent and targeted process that generates specialized chromatin domains, such as active genes, silent loci, or centromeres (Loyola and Almouzni 2004; Philpott et al. 2000). Hence, chaperones, along with nucleosome remodelers (not discussed here), establish chromatin plasticity and dynamics. Tuning by linker histones and their competitors A ninth histone (the linker histone, H1 or H5) joins the 2 nucleosomal DNA ends in a stem motif (Bednar et al. 1998; Hamiche et al. 1996b), reducing mononucleosome thermal breathing while, at the same time, facilitating conformational fluctuations between closed positive and negative states (Sivolob et al. in press; Sivolob and Prunell 2003). This last effect could, however, be hindered in the context of dense chromatin fibers, because the stacking of nucleosomes is favored by chromatin condensation upon linker histone binding (Bednar et al. 1998). Indeed, chromatosomal and nucleosomal templates respond similarly to torsional constraints (P. Recouvreux, private communication), suggesting that chromatin resilience should not change significantly upon linker histone binding. At the same time, chromatosome and nucleosome disruptions have been shown to occur in the same force range (Claudet et al. 2005), which is consistent with the high lability of H1 in vivo (Misteli et al. 2000). Several issues related to the linker histone are still highly debated, such as its exact binding mode on the nucleosome (Thomas 1999), its proposed role in nucleosome spacing (Woodcock et al. 2006), and its influence on chromatin fiber structure (Wong et al. 2007). Furthermore, as suggested earlier, the coupling between individual (at the nucleosome scale, as a nucleosome gate locker) and collective effects (at the chromatin fiber scale, as a chromatin condenser) provides H1 with versatile roles, for which mechanistic interpretations are not straightforward. H1 has been shown in vitro to hinder repair (Kysela et al. 2005) and transcription (Hannon et al. 1984; O’Neill et al. 1995; Shimamura et al. Published by NRC Research Press

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Transcription

Transcription-induced force RNA polymerase II (PolII), charged with the transcription of messenger (m)RNAs, is a 12 subunit (~500 kD) complex that unwinds the DNA double helix, polymerizes RNA, and proofreads the nascent transcript (Cramer 2004). Additional transcription factors and mediators can associate with PolII, to give a 60 protein (~3 MDa) complex (Boeger et al. 2005). Given the high molecular mass of the transcription machinery and the fact that it must drag the nascent RNA chain and prevent it from wrapping around the DNA template, DNA has been proposed to rotate around its axis during transcription, rather than polymerase turning around the template (Liu and Wang 1987). Some experimental evidence has supports this idea (Ostrander et al. 1990; ten HeggelerBordier et al. 1992; Wu et al. 1988). Probably the most straightforward argument in favor of a fixed polymerase is the proposal that transcription occurs in hundreds of discrete foci, called transcription factories (Cook 1999; Jackson et al. 1993), where chromatin is supposed to transiently migrate upon gene activation (see Fig. 2 and discussion later). Polymerases are among the most powerful molecular motors. First estimates of the force exerted by the Escherichia coli RNA polymerase was around 14 pN (Yin et al. 1995), a value that was soon upgraded to >25 pN (Wang et al. 1998), while the T7 DNA polymerase was shown to exert up to 40 pN on the DNA template (Wuite et al. 2000). Although all these measurements were made with prokaryotic enzymes, one can reasonably postulate that eukaryotic polymerases (potentially occasionally assisted by secondary molecular motors; see Actin and myosin in the nucleus) have enough power to pull the chromatin template through the viscous nucleus medium, unfold the supercoiled 30 nm fiber into a linear 10 nm nucleosomal array (less than 5 pN is needed for this transition (Cui and Bustamante 2000)), and disrupt nucleosomes that have been shown to be destabilized at ~20 pN (Bennink et al. 2001a; Brower-Toland et al. 2005, 2002; Claudet et al. 2005; Cui and Bustamante 2000; Gemmen et al. 2005; Mihardja et al. 2006; Pope et al. 2005). At the same time, forces around 10 pN have been shown to prevent nucleosome formation (Leuba et al. 2003), which means that once freed from the nucleosome, DNA should remain bare while under tension. Furthermore, the force required to disrupt nucleosomes is reduced under high ionic strength conditions (Gemmen et al. 2005; Mihardja et al. 2006), when histones are acetylated (Brower-Toland et al. 2005), and even more dramatically in the presence of histone chaperones (Bancaud et al. 2006). This means that plastic energy barriers can be reduced in vivo by a varied set of enzymes known for their dynamic interactions with chromatin, potentially lowering nucleosome disruption forces to the ~2 pN range expected from thermodynamic arguments (Marko and Siggia 1997).

Transcription in a chromatin context raises many questions, such as the way RNA polymerase progresses along the chromatin template, the subsequent fate of nucleosomes, and the dissipation of supercoiling constraints during the process (Lavelle 2007). Indeed, as DNA is rotated through the RNA polymerase, it follows a helical path (Harada et al. 2001), which means chromatin should undergo both traction and rotation during transcription elongation.

Transcription-induced torque Most single-molecule studies use topologically unconstrained DNA molecules. Recent experiments by Bancaud et al. (2006, 2007) have been done with torsionally constrained templates, closely mimicking the in vivo situation of topologically constrained chromatin fibers. Indeed, as mentioned before, translocation of DNA through a transcribing polymerase imposes a torque on the chromatin template.

1989). Catalyzed chromatosome movements could require specific nucleosome-remodeling factors (Maier et al. 2008). Last, H1 has many competitors for nucleosome binding in vivo, and could, thus, participate in a linker-protein network controlling nucleosomal DNA accessibility (Zlatanova et al. 2008). Among these competitors, high mobility group proteins are generally thought to give chromatin more plasticity and, in particular, to increase its transcriptional potential, although the mechanisms mediating these effects are not clear (Catez et al. 2004). Tuning by DNA functional conformational changes Single-molecule experiments on DNA have shown that the application of defined force and torque can drive different structural transitions (Sarkar et al. 2001). Hydrogen bonds and hydrophobic interactions, which stabilize B-DNA, involve free energies of only a few kT per bp. Indeed, a rather low torque of ~2 kT (remember 1 kT & 4 pN
nm) has proven sufficient to denature DNA (Strick et al. 1998). In vivo, DNA replication and transcription are linked to severe changes in the topological state of the double helix. As DNA is rotating inside the polymerase, positive and negative supercoiling is induced downstream and upstream, respectively (Liu and Wang 1987; see further discussion in Transcription). Transcriptionally generated torsion, rather than a mere waste product to be disposed of by topoisomerases, has instead recently been shown to propagate through the chromatin fiber and trigger local DNA alterations, detected as a regulatory signal by molecular partners (Kouzine et al. 2008). The free energy of negative supercoiling can locally melt the DNA helix, facilitating the opening of promoter regions, or drive, when reaching special DNA sequences, the formation of non-B (also called alternative or odd) DNA structures. These noncanonical structures include A-DNA (favored by reduced water content), Z-DNA (favored by GC and GT tracts), cruciforms (favored by inverted repeats), intramolecular triplexes and quadruplexes (H and H’-DNA, favored by mirror repeats; G-quartet, favored by G-rich sequences), slipped-strands (S-DNA, favored by direct repeats), and unpaired fragments. These transitions are thought to be functionally important in various biological functions, including gene expression, replication, recombination, and mutagenesis (Potaman and Sinden 2005; van Holde and Zlatanova 1994), and evidence is accumulating for their occurrence in vivo. Hence, noncanonical DNA and nucleosome structures may jointly appear upon and participate in transcription regulation (Lavelle 2008; Lavelle and Prunell 2007).

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Fig. 2. Force and torque occurrence in vivo: the case of transcription. This drawing is not intended to give a realistic picture of how transcription takes place in the nucleus, but only to schematically show how force and torque may participate in this process. Hence, for reasons of clarity, only nucleosomes and polymerases are represented, although numerous known (and probably as many unknown) factors participate in transcriptional initiation and elongation (including transcription factors, chromatin remodelers, histone chaperones). (a) A closed chromatin domain is tethered at the surface of a chromosome territory by some matrix attachment regions or insulator elements (Byrd and Corces 2003; Davie 1997). (b) Upon gene activation, the domain opens up. Two (potentially overlapping) mechanisms can assist this process: histone modifications can alter nucleosome charges and conformation, leading to a local decondensation and unfolding, potentially driven by chromatin fiber rotation (Lesne and Victor 2006); and ATP-dependent enzymes can actively modulate chromatin conformation and (or) apply some torsional constraints to the domain (Havas et al. 2000; Travers 1992). (c,d) Upon transcription, the domain further expands out of the chromosome territory. This could occur mainly through 2 distinct active processes. (c) Polymerase could, by itself, produce chromatin outlooping as a consequence of transcription elongation, performed in a factory located close to the border of the territory (Muller et al. 2007). (d) Alternatively, nuclear myosin I (NMI)/actin motor systems could drive the loop into a remote factory (Hofmann et al. 2006; Nunez et al. 2008), and polymerase would then pull chromatin through the factory (potentially shared by other transcribed genes from adjacent chromosome territories) during elongation. (e) Inside the factory, chromatin is pumped and screwed through the polymerase, undergoing both tension and torsion. Tension and positive supercoiling generated in front of the polymerase may help destabilize nucleosomes (Lavelle 2007). Negative supercoiling in the back may help reassemble nucleosomes and, at the same time, potentially induce some regulatory DNA secondary structures (Kouzine et al. 2008; Lavelle 2008). Interestingly, it has also been suggested that polymerase work is assisted by additional NMI/actin motors (de Lanerolle et al. 2005; Hofmann et al. 2006; Ye et al. 2008).

Strikingly, whereas a sufficient amount of tension disrupts nucleosomes, virtually any positive or negative supercoiling can be applied on a chromatin fiber without loss of nucleosomes (Bancaud et al. 2007). Such single-molecule torsion experiments, interpreted in the framework of the 3-state model of chromatin fiber (discussed earlier), provided a prediction of the torque as a function of supercoiling. Remarkably, this torque was found to vary, from 0 kT around the center of rotation of the fiber to a maximum of 0.75 kT at ~15 turns from the center, and remained at this

value while entering the plectoneme regime (Bancaud et al. 2006). At the same time, RNAP polymerase was found to exert a torque of at least 1.25 kT (potentially much higher) (Harada et al. 2001), which means that the torsional constraints imposed by the chromatin template will not hinder transcription. However, the 9 kT critical torque predicted for nucleosome torsional ejection (Sarkar and Marko 2001) may not be reached by the advancing polymerase, justifying the requirement of histone chaperones to assist transcription elongation in vivo (Workman 2006). Finally, the Published by NRC Research Press

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1.5 kT estimated torque necessary to trigger nucleosome– reversome transition suggests that reversome formation may help relax positive supercoiling ahead of the polymerase (Bancaud et al. 2007). Chromatin topological domains Human genes vary greatly in size, from 102 to more than 106 bp, with an average of ~30 kbp, which represents a significant fraction of the typical 5–200 kbp discrete chromatin topological domains thought to result from chromatin attachment to a fibrous structure, known as the nuclear matrix (Jackson et al. 1990), although this issue is controversial (Pederson 1998, 2000). Such an RNA-protein skeleton has been proposed to functionally organize the nuclear interior, mainly by providing some dynamic anchoring points for chromatin loops and transcription complexes (Davie 1997; Razin et al. 2007). Interestingly, matrix attachment regions have been shown to colocalize with insulators (Yusufzai and Felsenfeld 2004) and other protein complexes supposed to organize the eukaryotic genome into distinct domains (Bell et al. 2001). They have also been shown to allow decondensation of specific domains between its boundaries (Byrd and Corces 2003). However, limited information is available on the nature of topological domain boundaries that would present a barrier to rotation of DNA segments, as no clear evidence exists for any continuous fibrous skeleton in vivo. It is conceivable that protein-mediated anchoring of intracellular DNA to a nuclear membrane, skeleton, or other nuclear structure would provide such functional barriers within a living cell (Marenduzzo et al. 2007). Cook (1999) proposed that transcription factories, potentially linked with a nuclear skeleton, could participate in the partitioning of the genome. Interestingly, recent evidence of actin polymerization in the nucleus has enriched the palette of candidates for an intranuclear dynamic network participating in a structural scaffold to which the transcription machinery may be associated (Hofmann et al. 2006; Pestic-Dragovich et al. 2000; see Actin and myosin in the nucleus). Supercoiling during transcription Numerous studies have measured transcriptional velocity, giving values from 10 bases
s–1 to 100 bases
s–1, depending on the experimental conditions and the polymerase considered (Adelman et al. 2002; Darzacq et al. 2007; Harada et al. 2001; Shermoen and O’Farrell 1991; Thummel et al. 1990), with 70 bases
s–1 the maximum value measured for a eukaryotic polymerase in vivo (Darzacq et al. 2007). Questions abound concerning the fate of transient supercoiling produced by such highly processive tracking enzymes. Indeed, as already mentioned, transcription elongation is thought to generate nonconstrained positive supercoils ahead of the polymerase and negative supercoils behind it (Liu and Wang 1987). Note that this dynamic nonconstrained supercoiling adds to the nonconstrained positive supercoiling induced within the topologically closed chromatin domain to compensate constrained negative supercoils produced by the promoter unwinding (Revyakin et al. 2004). However, this initiation-induced supercoiling (roughly 1.2 turns) is negligible in comparison with the much higher elongationinduced supercoiling. Indeed, if translocation proceeds without slippage, which has proven to be the case for bac-

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terial RNA polymerase (Harada et al. 2001), 1 negative rotation should be generated for every 10.5 bp covered, which represents ~2850 turns for a typical 30 kbp gene! It is noteworthy that high levels of transcription driven by multiple polymerases in a tandem array would not produce higher levels of supercoiling (neglecting the previously mentioned supercoiling component due to promoter unwinding). Indeed, positive and negative supercoiling generated by polymerase translocation should partially neutralize between successive polymerases within the array. From a kinetic point of view, the amount of supercoiling generated by a polymerase translocating at ~10–100 bp
s–1 should be roughly 1–10 supercoils
s–1, potentially exceeding the relaxation capability of DNA topoisomerases, as measured from bulk in vitro relaxation kinetic studies (Osheroff et al. 1983), as well as single-molecule experiments on a naked DNA template (Strick et al. 2000). Furthermore, the presence of nucleosomes in vivo has been shown to alter the way topoisomerases relax topological constraints (Di Felice et al. 2008; Salceda et al. 2006), and localized supercoiling may escape relaxation to mediate topological couplings between DNA transactions (Wang and Droge 1996) and influence transcription regulation (Kouzine et al. 2008; see following section). Interestingly, PolII has been shown in vivo to slow down from 70 bases
s–1 to 16 bases
s–1 when relaxation by topoisomerase (topo)I is inhibited upon treatment with camptothecin (Darzacq et al. 2007), presumably as a consequence of residual torsional stress, as suggested by previous studies (Collins et al. 2001). The previous results suggest a scheme to explain how transcription elongation could benefit from the high torsional resilience of chromatin. This feature may indeed serve to cushion the supercoiling waves generated by polymerases during transcription. At the same time, chromatin fiber resilience could dampen the driving torque produced by polymerase translocation. This would partly explain the low activity of topoI, a torque-sensitive topoisomerase (Koster et al. 2005), on a nucleosomal template (Salceda et al. 2006) while, at the same time, chromatin might favor the DNA transport activity of topoII by increasing the juxtaposition probability of DNA segments and facilitating the nucleosome entry and exit DNAs to shift from a closed negative state to a closed positive one (Lavelle 2007; Salceda et al. 2006). One can, moreover, speculate that topoI should mainly relax negative supercoiling produced in the wake of the polymerase and present as negative torsional constraint in linker DNA (Kouzine et al. 2008). TopoII, however, would be more efficient in relaxing positive supercoiling produced in front of the polymerase and present as positive writhe, since no sink for torsional stress, such as melting, exists in this case (Lavelle 2008). However, in regions where the transcription rate is high, nucleosomes may be depleted by frequent polymerase passage and DNA-pulling forces exerted by polymerases, which may also prevent the local formation of supercoils. Helical tension would then deform DNA, mostly by twist, allowing topoI to be more efficient than topoII. This scheme is consistent with the preferential localization of topoI in highly transcribed regions (see Lavelle 2007 and references therein). Interestingly, a recent study showed that a rapid (prior to PolII passage) loss of nucleosomes occurs at Drosophila melaPublished by NRC Research Press

Lavelle

nogaster Hsp70 locus after heat shock induction (Petesch and Lis 2008), where topoI had been formerly shown to be concentrated (Gilmour et al. 1986), whereas topoII was mostly found in flanking regions (Udvardy et al. 1985). However, although new studies regularly stress the crucial role of topoisomerases for transcription elongation (see, for instance, Muller et al. 2007), it is still quite difficult to get an unambiguous mechanistic interpretation, since these enzymes have multiple roles in the nucleus (Borde and Duguet 1998; Nitiss 1998). DNA as a mechanosensor participating in transcriptional regulation Due to the viscous drag on DNA and chromatin fiber in the nuclear medium, transient torques can develop even in the absence of DNA tethering (Nelson 1999). Indeed, transcriptionally generated torque has been shown to be sufficient to melt susceptible sequences upstream of active promoter in topologically open linear DNA in vitro (Kouzine et al. 2004). This effect should, at the same time, generate some transient unconstrained supercoils. However, the torsional wave will propagate much more rapidly than the advancing polymerase on DNA (Sarkar and Marko 2001), and presumably also on chromatin (J.-M. Victor, private communication). Topoisomerases do not instantaneously relax these supercoils (Kouzine et al. 2008; Salceda et al. 2006). The preservation of some superhelical tension in active chromatin was suggested to maintain a transcriptionpoised chromatin state (Villeponteau et al. 1984), and could also help stabilize odd DNA structures (Kouzine et al. 2008). Indeed, Kouzine and coworkers (2008) recently measured the transcription-generated supercoiling in chromatin in vivo; before slowly decaying, the superhelical tension was able to trigger non-B-DNA structure in a specific supercoiling-sensing sequence within a linker, located 6 nucleosomes upstream of the promoters. This non-B-DNA structure, in turn, recruited 2 transcriptional factors essential for the expression of the gene (Liu et al. 2006). Therefore, in addition to providing a cushion to transcription-induced supercoiling waves, chromatin may act as both a clutch and a driving belt to modulate transmission of those waves for the dynamic control of gene expression (Lavelle 2008). Alternatively, secondary structures in DNA, by absorbing negative supercoiling constraints in the wake of RNA polymerase, could hinder DNA melting and, thus, lower pausing and instability caused by RNA/DNA hybridization (R-loop) between the RNA transcript and the template DNA strand (Drolet 2006; Li and Manley 2005).

Large-scale chromatin organization With an estimated 60%–70% water fraction in the nucleus, the nuclear interior may be considered as a soft aqueous gel (Rowat et al. 2006) in which macromolecular crowding, with up to 100 mg
mL–1 macromolecular concentration, induces the formation of subnuclear structures (Hancock 2004). In this medium, interphasic chromosomes appear to be compartmentalized into tightly compacted chromosome territories separated by less compact interchromatin domains (Branco and Pombo 2007; Cremer et al. 2006). It has been proposed that interchromatin domains spread into

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chromosome territories through channels, to form a chromatin architecture reminiscent of a sponge (Cremer and Cremer 2001; Visser et al. 2000), with transcription sites scattered throughout chromosome territories (Verschure et al. 1999). The position of genomic regions inside this particular architecture is believed to participate in cell differentiation and gene expression regulation (Francastel et al. 2000; Lamond and Earnshaw 1998; O’Brien et al. 2003). Chromatin outlooping Several studies have shown that genomic regions dynamically expand from the surface of chromosome territories into interchromatin domains upon transcription (Volpi et al. 2000; Williams 2003) (Fig. 2). This process requires local and specific chromatin decondensation (opening) and extrusion (outlooping), and has been suggested to be influenced by DNA topological constraints (Lesne and Victor 2006). It is also believed to drive targeted genes into functional nuclear compartments, such as transcription factories (Bartlett et al. 2006; Jackson et al. 1993; Osborne et al. 2004), which have been shown to exist as independent nuclear subcompartments, rather than being simple accumulations of PolII on transcribed genes (Mitchell and Fraser 2008). Most genes move in and out of these sites, depending on their transcription status, whereas highly expressed genes seem to be constantly associated with a factory (Osborne et al. 2004). Outlooping generally correlates with decondensation of the chromatin (Chambeyron and Bickmore 2004), although these 2 mechanisms can also occur independently, which means outlooping of a locus is not merely a consequence of chromatin decondensation (Morey et al. 2007). Rather, during outlooping, a limited domain of chromatin fiber probably undergoes a structural transition, from an inactive structure to an active one in which nucleosomes are no longer stacked upon each other. Such coupling between compaction and transcription states of a locus was suggested a long time ago (Andersson et al. 1982). From a mechanistic point of view, chromatin 30 nm fiber decondensation has been proposed to correlate with torsion of the fiber axis (Barbi et al. 2005), suggesting a physical decondensation mechanism that would rely on topological constraints (Lesne and Victor 2006), potentially with the assistance of some ATP-dependent enzymes, such as helicases or remodeling factors (Havas et al. 2000; Travers 1992), due to their ability to twist DNA when translocating on it (Lia et al. 2006). Such topologically driven loop condensation and decondensation provides a convenient mechanism linking the nucleosome state in the loop to the overall chromatin domain structure. The functional significance of higher-order chromatin structure and dynamics in regulating individual gene expression, along with its control by various biochemical events (covalent histone modifications, nucleosome remodeling), remains speculative. Chromatin folding should depend dramatically on the state of its elementary components (see Chromatin physical tuning). Consistent with this view, histone marks that are associated with active transcription are still present on decondensed chromatin after transcription, keeping the domain in an open state that would facilitate reinitiation (Muller et al. 2007). Looking at it more closely, chromatin seems, in fact, to undergo 2 kinds of energy-dependent motions (Soutoglou Published by NRC Research Press

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and Misteli 2007): short-range random motions, probably resulting from continuous opening and closing events of chromatin by ATP-dependent remodeling machines (Figs. 2a and 2b); and long-range directed motions, produced by an ATPdependent molecular motor (Figs. 2c and 2d). Interestingly, recent data involving constrained diffusion dependent on actin–myosin cooperative interaction dramatically renewed our vision of the mechanisms underlying long-range chromatin movements (Chuang et al. 2006). Actin and myosin in the nucleus As already discussed, the issue of intranuclear structure promoted by macromolecular filaments, which would somehow mimic the cytoplasmic architecture, has been a matter of debate for more than 50 years (Pederson 2000). Whereas the presence of a lamina structure at the nuclear periphery has been clearly demonstrated (Aebi et al. 1986; Paddy et al. 1990), there is less evidence for an internal structural network, although some data have pointed out stable structures formed by intranuclear lamins (Bridger et al. 2007; Moir et al. 2000). Remarkably, the characterization of nuclear actin brought a new twist on the issue. Actin filaments are well known for their role in defining cell shape and motility, intracellular transport, and contraction of muscle cells. Under physiological salt conditions and above a critical concentration, monomers (globular- or G-actin) polymerize into filaments (filamentous- or F-actin); this process, along with interaction with myosin, is involved in virtually all known functions of actin in the cytoplasm. The existence of nuclear actin was reported 40 years ago (Lane 1969), and it has since been confirmed by many studies establishing diverse functions of actin in the nucleus (Bettinger et al. 2004). Only recently was a polymeric form of actin in the nucleus clearly demonstrated (Hofmann and de Lanerolle 2006; McDonald et al. 2006), making actin a fundamental actor in nuclear roles that include chromatin remodeling, gene transcription, transport, and maintenance of nuclear structure (Percipalle 2007). Interest in intranuclear motor functions was similarly stimulated by the discovery of a nuclear isoform of myosin, nuclear myosin I (NMI) (Nowak et al. 1997). Subsequently, NMI was shown to be associated with RNA polymerases I (Fomproix and Percipalle 2004), II (Hofmann et al. 2004), and III (Hu et al. 2004), and has also been shown to interact with many chromatin modifying and remodeling enzymes (Chen and Shen 2007; Olave et al. 2002). Because all myosins are actin-activated ATPases, an acto-myosin molecular motor is supposed to act in the nucleus (Philimonenko et al. 2004). Indeed, several recent studies have observed intranuclear long-range chromosome movements and have demonstrated the role for an actin–myosin complex in the active and directed translocation of chromatin domains (Chuang et al. 2006; Dundr et al. 2007; Nunez et al. 2008) (Fig. 2). These studies have clarified how an inactive chromosome region may translocate, upon activation, toward the interior of the nucleus (Chuang et al. 2006) or toward particular nuclear organelles, such as Cajal bodies (Dundr et al. 2007), and how long-distance chromosomal interactions may take place (Nunez et al. 2008). These movements occurred at velocities ranging from 0.1 to 1.0 mm
min–1 and over distances of 1–5 mm. Since motors have to be anchored to generate

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force, it is tempting to suggest that NMI interacts with some type of actin polymer, although these nuclear polymers are different from cytoplasmic filaments (McDonald et al. 2006). Alternatively, actin has been proposed to relax chromatin structure, allowing an activated locus to loop outside of the chromosome domain and reach neighboring compartments (Carmo-Fonseca 2007), but the mechanism underpinning this possibility is not clear. Moreover, polymeric actin has recently been shown to cooperate with NMI to power PolI transcription (Ye et al. 2008), confirming a hypothesis proposed 3 years earlier (de Lanerolle et al. 2005). But again, no clear mechanism has emerged for this process. Taken together, these studies provide a glimpse of the growing picture of the multifaceted role of these former cytoplasmic proteins.

Conclusion Chromatin is a polymorphic substrate, comprising mostly DNA–histone structures driven by more or less specific interactions and influenced by DNA topology. Time has come to transpose the mechanistic approach of DNA, considered as a molecular spring — which has proven valuable for the study of many biological processes, including DNA–protein recognition, nucleosome positioning, and transcription initiation — to the next and more physiological structural level, that is, the chromatin fiber, which may be regarded as a complex and polymorphic functional supramolecular metaspring. In this quest of mechanistic understanding of transcriptional regulation, a detailed description of the nuclear architecture, with the function of its subcompartments, and of the genome organization is obviously required (Branco and Pombo 2007; Dundr and Misteli 2001; Pombo and Branco 2007; Spector 2003, 2006), along with the extent to which nuclear structural proteins (lamin, actin) influence high-order chromatin structure and tissue-specific gene expression. In parallel, enzymatic studies (relaxation, transcription) should now be performed on single chromatin templates. The accumulation of new quantitative data and molecular models provided by these complementary approaches should help to drive us toward a more consistent and biologically relevant mechanistic view of genetic expression within the next years. I hope this short overview will stimulate further investigation in this exciting interdisciplinary field.

Acknowledgements I thank Annick Lesne, Se´bastien Neukirch, Jean-Marc Victor, and Aure´lien Bancaud for helpful discussions, Keir Neuman for proofreading of the manuscript, Hua Wong for considerable help in making Fig. 2, and the 2 anonymous referees for fruitful comments on the manuscript.

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